Abstract
Tumor metastasis remains the leading cause of mortality among cancer
patients. Addressing this challenge necessitates the development of
effective strategies for targeted drug delivery and therapy. Given that
metastatic lesions are primarily driven by highly aggressive tumor cell
subpopulations, in‐depth study of these cells and further guiding
design of targeted therapeutics, play deterministic roles in metastasis
inhibition. Herein, a nano‐electro‐platform is shown that enables
non‐invasive screening of aggressive cell subpopulations from
heterogeneous tumor samples. Single‐cell sequencing further reveals
immune evasion pathways associated with their aggressive behavior.
Targeting the screened aggressive cells, the platform implements a
unique nanopore‐focused electric field, which genetically remodels the
cells to generate extracellular vesicles (EVs) with significantly
enhanced tumor‐targeting and therapeutic capabilities. The engineered
EVs effectively activate macrophages and T cells, leading to robust
tumor cell elimination and metastasis inhibition in lung cancer
metastasis models. These highlight a versatile, multidisciplinary
technique adopting a new path toward deep understanding and treating
metastasis.
Keywords: electroporation, extracellular vesicles, immunotherapy,
nano‐chip, tumor metastasis
__________________________________________________________________
The nano‐electro‐platform achieves non‐destructive evolutionary
screening and electro‐remodeling of tumor cells to generate
extracellular vesicles with enhanced tumor‐targeting and therapeutic
functions. These extracellular vesicles demonstrate remarkable efficacy
in activating both macrophages and T cells, thereby orchestrating a
potent immune response that results in the elimination of tumor cells
and the suppression of metastatic progression within lung cancer
metastasis models.
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1. Introduction
Tumor metastasis is the leading cause of mortality in cancer patients,
making mechanism research and effective treatment imperative.^[ [64]^1
, [65]^2 ^] Due to the high heterogeneity of tumors, metastatic foci
are predominantly formed by a subpopulation of aggressive cells
behaving enhanced migration and invasiveness. Screening the aggressive
subgroup for decoding the underlying mechanism brings about important
hints for inhibiting tumor metastasis.^[ [66]^3 , [67]^4 , [68]^5 ^]
However, current screening methods including flow cytometry depend on a
single or a low number of biomarkers, falling short of precise
differentiation of the subgroup orchestrated by a variety of genetic
regulation.^[ [69]^6 , [70]^7 ^] Furthermore, based on a limited
understanding of the subgroup, traditional therapeutics (i.e.,
chemotherapy drugs and immunotherapy inhibitors) for inhibiting tumor
metastasis face challenges to achieving high efficiency in targeting
and precise elimination.^[ [71]^8 , [72]^9 , [73]^10 , [74]^11 ^]
Technologies that enable precise screening of the highly migratory and
invasive subgroups while facilitating the development of targeted drugs
have been sought after for the long term yet remained wide open to
tumor metastasis inhibition.
Here, we developed a nanoplatform for evolutionary Screening and
Electro‐remodeling of tumor cells to generate Extracellular vesicles
with enhanced tumor‐targeting and therapeutic functions (“SEE”
platform). The platform first adopts a 3D cell‐culture module for
mimicking in vivo invasion of cancer cells, to screen aggressive cell
subpopulation (AG cells) from the patient's tumor tissue, which
bypasses the limit of biomarkers. Single‐cell sequencing next revealed
key genes, including carbonic anhydrase IX (CA9) and lactamase beta 2
(LACTB2) related to immune escape, determining the migratory and
invasive variances between AG cells and their non‐aggressive
counterparts. Facilitated by the SEE platform, new findings relevant to
AG cells further indicate that their released EVs present significantly
enhanced capacity to targeting tumors, encouraging a cell‐remodeling
strategy for metastasis inhibition by adopting the engineered EVs. To
this aim, the SEE platform adopts a nanopore‐focused electric field
that not only achieves safe, efficient intracellular delivery of
therapeutic molecules for remodeling AG cells but also triggers the
remodeled cells to produce therapeutic‐EVs (SEE‐EVs) that efficiently
transport into targeted tumor cells. For in vivo application toward
metastasis treatment, the SEE platform was applied for remodeling AG
cells from lung cancer patients by transfecting immunotherapy
promoters, Interferon‐γ (IFN‐γ) plasmid, and Programmed cell death
ligand 1(PD‐L1) siRNA. Systematic investigations based on both
cell‐derived xenograft (CDX) and patient‐derived xenograft (PDX) models
validated the significantly enhanced therapeutic effects of the SEE‐EVs
to host immune system, regaining its abilities in identifying and
eliminating tumor cells. These highlight a versatile, multidisciplinary
nano‐electro‐platform adopting a unique path toward studying and
treating tumor metastasis.
2. Results
2.1. Working Principle of the SEE Platform
The SEE platform consists of an evolutionary screening (ES) module and
an electro‐remodeling (ER) module (Figure [75]1A–C; Figures [76]S1A–F
and [77]S2, Supporting Information). The ES module was designed to
non‐destructively isolate AG cell subpopulation from patient‐derived
tumor tissues. To achieve this, we developed a culture chamber with a
flat base and gradient sidewall to simulate the in vivo migration and
invasiveness of AG cells from the primary tumor site (flat region) to
metastatic niches (gradient region). The gradient region creates a 3D
spatial barrier for cell differentiation.^[ [78]^12 ^] Due to
significant variances on migration and invasiveness, AG cells can be
isolated from the gradient region. The subgroup with low migration,
named “ambitionless (AM) cells”, remains in the flat region.
Figure 1.
Figure 1
[79]Open in a new tab
Principle of the SEE platform for metastasis inhibition. A) Acquisition
of tumor samples from cancer patient. B) Layout of the SEE platform,
including the evolutionary screening (ES) module (left) and
electro‐remodeling (ER) module (right). AG cell, aggressive cell. AM
cell, ambitionless cell. SEE‐EVs, EVs with tumor‐targeting and
therapeutic functions. C) Structural diagram of the SEE platform. D)
Schematic diagram illustrating the aggregation of SEE‐EVs inside the
tumor. E) Schematic diagram of SEE‐EVs inhibiting tumor metastasis. F)
Selection of the slope for evolutionary screening. G) Scratch test to
verify the enhanced migration capacity of evolutionarily screened
cells. H) 3D invasion assay to verify the increased aggressiveness of
the evolutionarily screened cells. The data shown in F‐H were derived
from three independent experiments and were presented as mean ± SD; a
two‐sided Student's t‐test was used for comparisons (F‐H). p < 0.05 was
considered statistically significant.
Further study on the AG cells isolated by the SEE platform reveals that
the EVs released from them have significantly enhanced performance in
tumor‐targeting, which encourages adopting the AG‐derived EVs as drug
carriers for tumor metastasis inhibition. For developing the engineered
therapeutic‐EVs (SEE‐EVs), the ER module is designed on the platform,
where a nanopore layer is assembled, forming a unique configuration
that generates a focused electric field to safely electroporate the AG
cells (cultured on the nanopore layer) while rapidly transporting
therapeutic factors into cells. Facilitated by the nanopore‐focused
electric field (NEF), the transfected AG cells significantly produce
the SEE‐EVs that carry the therapeutic factors.
By using lung cancer as a proof‐of‐concept model, IFN‐γ and PD‐L1 siRNA
were applied as therapeutic molecules for restoring the tumor‐killing
function of immune cells, including macrophages and T cells, which play
crucial roles in eliminating tumor cells. The precursor of IFN‐γ (i.e.,
IFN‐γ mRNA) was formed by the transcription of IFN‐γ plasmid following
its delivery into AG cells and then co‐encapsulated with the delivered
PD‐L1 siRNA into the EVs secreted by AG cells. After being reinfused
back into the tumor model, the SEE‐EVs efficiently propagate to tumor
cells and trigger the intracellular release of IFN‐γ mRNA and PD‐L1
siRNA (Figure [80]1D). The intracellular IFN‐γ secretion enhanced by
IFN‐γ mRNA led to the polarization of macrophages from the M2 phenotype
(which promotes tumor growth) to the M1 phenotype (which exerts
tumoricidal effects). Meanwhile, the reduced PD‐L1 expression, as
inhibited by PD‐L1 siRNA, reactivated the T cells, encompassing both
CD4^+ T cells with the ability to recognize tumor cells and CD8^+ T
cells for subsequent tumor elimination (Figure [81]1E). Through the
dual killing effect (by activating macrophages and T cells), the
SEE‐EVs enable effective inhibition of tumor metastasis.
2.2. The SEE Platform for Screening Aggressive Cells
To validate the SEE platform for screening AG cells, we focus on lung
cancer in this work, in recognition of its high prevalence and
metastatic potential.^[ [82]^13 , [83]^14 ^] The screening results of
lung cancer cells (A549 and H1975) demonstrate that migratory distance
decreases significantly at 30° as compared to 20°, while slopes
≥50°(50°, 60°, 70°, and 80°) show no statistically significant
differences in migration distance (Figure [84]1F; Figure [85]S3A,B,
Supporting Information). Accordingly, the slope of 50° was selected for
screening AG cells. To further validate the migratory and invasive
capacities of AG cells, we compared the cells isolated from 50° to
their counterparts remained on the flat region. The results of the
scratch assay show a higher confluence of the cells from the slope
(Figure [86]1G; Figure [87]S4A,B, Supporting Information). In addition,
the 3D invasion assay suggests an increased rate of invasive cells in
the slope group (Figure [88]1H; Figure [89]S5A–E, Supporting
Information), revealing that AG cells exhibit significantly enhanced
migration and invasiveness capacities. Additionally, the cells without
screening (Control group) exhibit complex features of both the AG cells
and AM cells, which implies the heterogeneity of cell behaviors.
To further investigate the mechanisms underlying the enhanced migration
and invasiveness capabilities of AG cells, we performed transcriptomic
profiling of the AG cells and AM cells. The results highlight 41
upregulated genes and 15 downregulated genes in AG cells of both lung
cancer cell lines (Figure [90]SE1A–D, Supporting Information). Among
the changed genes, most are closely related to the immune escape of
tumors through the inhibition of macrophages and T cells, such as CA9
^[ [91]^15 , [92]^16 ^] and LACTB2.^[ [93]^17 , [94]^18 ^] These
results indicate that the enhanced capabilities of AG cells are mainly
derived from the regulation of the immune microenvironment.
Transcriptomic profiling further reveals hundreds of personalized genes
in determining the aggressive behaviors in the two lung cancer cell
lines (Figures [95]SE1E–J and [96]S6A–F, Supporting Information). These
data confirm the genetic heterogeneity of cells while indicating the
independence of biomarker limitations by adopting the SEE platform for
screening AG cells.
Taken together, the SEE platform with a slope‐shaped ES module
effectively screens aggressive cell subpopulations with enhanced
migration and invasiveness from lung cancer cells. Further
investigations identify key genes governing the aggressive behaviors of
AG cells, primarily implicated in tumor immune evasion. These findings
elucidate potential tumor metastasis mechanisms and offer insights for
subsequent drug development strategies.
2.3. AG Cell‐Derived EVs with Enhanced Tumor‐Targeting Ability
Recent studies have indicated enhanced tumor‐targeting potentials of
cancer cell‐derived EVs, along with additional advantages of high
tissue penetration and low immunogenicity.^[ [97]^19 , [98]^20 ^] These
encouraged us to investigate whether the EVs derived from the
SEE‐screened AG cells exhibit functional strengths as a delivery system
for metastasis inhibition. For this purpose, EVs from the cells
isolated from the SEE platform with 30° (EVs‐30°), 50° slopes
(EVs‐50°), and the cells without screening (EVs‐0°) were compared in
parallel. The results show no significant differences on morphology
(Figure [99]2A; Figure [100]S7A, Supporting Information), size
distribution (Figure [101]2B; Figure [102]S7B,C, Supporting
Information), and expression of EV markers (CD63, CD9, TSG101)
(Figure [103]2C; Figure [104]S7D, Supporting Information). However, the
cellular internalization efficiency of EVs from the cells isolated from
the SEE platform is significantly increased, in direct proportional to
the screening slope gradient (Figure [105]2D,E; Figure [106]SE2A–D,
Supporting Information). Furthermore, parental cells co‐cultured with
EVs‐50° resulted in an 8.95‐fold increase in EVs uptake, as compared to
that of non‐parental cells (Figure [107]2F,G; Figure [108]SE2E–H,
Supporting Information).
Figure 2.
Figure 2
[109]Open in a new tab
Evolutionary screening enhanced the targeting of EVs to tumor cells. A)
TEM images of EVs (scale bars = 50 nm). 0°, 30°and 50° represents EVs
derived from cells screened at the slopes of 0° (EVs‐0°), 30° (EVs‐30°)
and 50° (EVs‐50°), respectively. B) NTA analysis of EVs. C) Western
blot analysis of marker proteins of EVs (CD63, CD9, and TSG101). D,E)
Fluorescence images (D) and quantitative analysis (E) of the uptake of
EVs by parental A549 cells (scale bars = 50 µm). Red: PKH26‐labeled
EVs; Blue: DAPI‐stained cell nuclei. F,G) Fluorescence images (F) and
quantitative analysis (G) of the uptake of EVs‐50° by different cell
types (scale bars = 25 µm). Red: PKH26‐labeled EVs; Blue: DAPI‐stained
cell nuclei. H,I) In vivo distribution (H) and quantitative analysis
(I) of EVs in A549 tumor‐bearing BALB/c nude mice at different time
points after intravenous injection. J,K) Ex vivo fluorescence images
(J) and quantitative analysis (K) of EVs in the main organs from A549
tumor‐bearing BALB/c nude mice. L) Frozen sections of the liver and
tumor. The nuclei and EVs are indicted by blue and red fluorescence
(marked with green circles), respectively (scale bars = 25 µm). The
data shown in E, G, and K were derived from three independent
experiments and were presented as mean ± SD; a two‐sided Student's
t‐test was used for comparisons (E,G,K). p < 0.05 was considered
statistically significant.
For further validation of the uptake difference in vivo, we established
murine tumor models and injected EVs‐0°, EVs‐30°, and EVs‐50°
respectively via the tail vein. Notably, EVs‐30° and EVs‐50° exhibit
significantly higher accumulation in tumor tissues compared to EVs‐0°,
with EVs‐50° demonstrating the most efficient tumor‐targeting capacity.
By contrast, EVs‐0° predominantly accumulate in the liver
(Figure [110]2H–L; Figure [111]SE2I–R, Supporting Information). These
results demonstrate the enhanced tumor‐targeting of AG cell‐derived
EVs.
To investigate the mechanisms underlying the tumor‐targeting of EVs
derived from AG cells (EVs‐AG), we performed proteomic profiling on
EVs‐AG and EVs derived from AM cells (EVs‐AM). The results show that
EVs‐AG in both A549 and H1975 cell lines commonly exhibit 9 upregulated
proteins and 14 downregulated proteins (Figure [112]3A–C). The
differentially expressed proteins are mainly involved in protein
recognition and cellular binding, with predominant localization in the
extracellular region, vesicles, and extracellular space. In terms of
biological processes, the identified proteins are mainly linked to
metabolic regulation (Figure [113]3D–G). Additionally, over 40% of the
differentially expressed proteins participate in biological activities
through secretory mechanisms, suggesting a key role in extracellular
signaling and intercellular communication. (Figure [114]S8A,B,
Supporting Information).
Figure 3.
Figure 3
[115]Open in a new tab
Differential proteomic analysis of EVs derived from evolutionarily
screened cells. A,B) Venn diagram of up‐(A) and downregulated (B)
proteins in EVs from AG cells (EV@cell‐AG) versus EVs from AM cells
(EV@cell‐AM). Coloring based on cluster identifiers. The left circles
represent the number of up‐regulated (A) and down‐regulated (B)
proteins in EV@A549‐AG versus EV@A549‐AM, respectively. The right
circles represent the number of up‐regulated (A) and down‐regulated (B)
proteins in EV@H1975‐AG versus EV@H1975‐AM, respectively. Overlapping
segments represent the number of genes that are jointly up‐regulated
(A) and down‐regulated (B) in the two groups. C) The up‐(orange) and
downregulated (green) proteins in EV@cell‐AG versus EV@cell‐AM. D,E) GO
(D) and KEGG (E) pathway enrichment analysis of differently expressed
proteins between EV@A549‐AM and EV@A549‐AG. F,G) GO (F) and KEGG (G)
pathway enrichment analysis of differently expressed proteins between
EV@H1975‐AM and EV@H1975‐AG. The screening criterion was
fold‐change > 1.5‐fold and p < 0.05 were considered statistically
significant. P‐values were calculated using fisher's exact test with
the hypergeometric algorithm and adjusted using the Benjamini‐Hochberg
method for multiple tests (D–G).
Taken together, the EVs derived from the SEE‐isolated AG cells exhibit
significantly enhanced tumor‐targeting efficiency, showing potentials
as a delivery system for targeted metastasis inhibition. The study of
proteomic profiling reveals that their enhanced affinity toward tumor
cells mainly derives from the regulation of proteins related to
cellular recognition and binding.
2.4. The SEE‐Mediated AG Cell Electro‐Remodeling for Engineering Therapeutic
EVs
Encouraged by the enhanced tumor‐targeting capability of the AG
cell‐derived EVs, we applied the SEE platform for further engineering
AG cells to produce therapeutic EVs. To achieve this, the SEE platform
adopts the ER module that was designed for transfecting AG cells with
therapeutic factors by a nanopore‐focused electric field (NEF). The
simulation of the electric field distribution demonstrates that the NEF
reaches transmembrane potential at a low voltage, enabling transient
and reversible opening of the cell membrane.^[ [116]^21 , [117]^22 ,
[118]^23 ^] Meanwhile, charged molecules could be directionally
transported into the cell (Figure [119]S9A–D, Supporting Information).
Compared to traditional cell‐engineering methods, including liposome
(Lipo)^[ [120]^24 ^] and bulk electroporation (BEP),^[ [121]^25 ^] the
SEE platform demonstrates higher cell viability (up to 96%) and
delivery efficiency (up to 93%) (Figure [122]4A; Figure [123]S10A–H,
Supporting Information). Furthermore, the SEE platform significantly
enhances EVs production, achieving a 2.49‐fold increase compared to
Lipo and a 1.60‐fold increase relative to BEP (Figure [124]4B; Figure
[125]S11A–H, Supporting Information). This may be attributed to the
NEF‐based delivery on the SEE platform, which maintains high cell
viability while upregulating intracellular molecules associated with EV
production.^[ [126]^26 , [127]^27 ^]
Figure 4.
Figure 4
[128]Open in a new tab
Characterization of EVs endowed with therapeutic properties. A) Cell
viability and efficiency of the delivery of GFP plasmid into A549 cells
using Lipo, BEP, and NEF, respectively. B) Total protein of EVs from
A549 cells after the delivery using Lipo, BEP, and NEF, respectively (n
= 6). C,D) Content of IFN‐γ mRNA (C) and PD‐L1 siRNA (D) in A549 cells
after delivery using NEF. Control: untreated cells; IFN‐γ: cells loaded
with the IFN‐γ plasmid; PD‐L1: cells loaded with PD‐L1 siRNA;
IFN‐γ+PD‐L1: cells loaded with both the IFN‐γ plasmid and PD‐L1 siRNA.
E) TEM images of EVs (scale bars = 50 nm). (E–I) Control: EVs derived
from untreated cells; IFN‐γ: EVs derived from cells loaded with the
IFN‐γ plasmid; PD‐L1: EVs derived from cells loaded with PD‐L1 siRNA;
IFN‐γ+PD‐L1: EVs derived from cells loaded with both the IFN‐γ plasmid
and PD‐L1 siRNA. F) NTA analysis of EVs. G) Western blot analysis of
marker proteins of EVs (CD63, CD9, and TSG101). H,I) Content of IFN‐γ
mRNA (H) and PD‐L1 siRNA (I) in EVs. J,K) Content of IFN‐γ mRNA (J) and
PD‐L1 mRNA (K) in parental cells after incubation with EVs. (J–N)
Control: parental cells after incubation with EVs derived from
untreated cells; IFN‐γ: parental cells after incubation with EVs
derived from cells that were loaded with the IFN‐γ plasmid; PD‐L1:
parental cells after incubation with EVs derived from cells that were
loaded with PD‐L1 siRNA; IFN‐γ+PD‐L1: parental cells after incubation
with EVs derived from cells that were loaded with both the IFN‐γ
plasmid and PD‐L1 siRNA. L) Western blot analysis of IFN‐γ and PD‐L1
protein levels in parental cells after incubation with EVs. M,N)
Quantitative analysis of IFN‐γ (M) and PD‐L1 protein levels (N) in
parental cells after incubation with EVs. The data were derived from
three independent experiments unless otherwise stated and were
presented as mean ± SD; a two‐sided Student's t‐test was used for
comparisons (A–D, H–K, M and N). p < 0.05 was considered statistically
significant.
In terms of the therapeutic molecules, given the critical link between
immune escape gene networks and AG cell invasiveness,^[ [129]^28 ,
[130]^29 , [131]^30 ^] we selected a therapeutic strategy targeting two
pathways: macrophages^[ [132]^31 , [133]^32 ^] and T cells.^[ [134]^33
, [135]^34 , [136]^35 ^] The engineering process comprises three steps,
including 1) Intracellular delivery: IFN‐γ plasmid and PD‐L1 siRNA are
delivered into screened AG cells to increase intracellular levels of
IFN‐γ mRNA and PD‐L1 siRNA; 2) EV secretion: AG cells secret EVs loaded
with IFN‐γ mRNA and PD‐L1 siRNA, termed “SEE‐EVs”; 3) Therapeutic
delivery: leveraging the high tumor‐targeting capability of SEE‐EVs in
vivo, IFN‐γ mRNA and PD‐L1 siRNA are transported into tumor cells,
resulting in enhanced IFN‐γ secretion and downregulation of PD‐L1
expression.
We first validated whether these cells were genetically engineered by
quantifying RNA levels. The significant increase in intracellular IFN‐γ
mRNA (>8.43‐fold) and PD‐L1 siRNA (>19.32‐fold) confirms the
implementation of cell engineering (Figure [137]4C,D; Figure
[138]SE3A–D, Supporting Information). Furthermore, in cells receiving
exclusive delivery of either IFN‐γ plasmid or PD‐L1 siRNA, the
upregulated molecules correspond to the delivered payloads. We then
collected EVs derived from the engineered cells and characterized their
properties. The results show no significant differences in morphology
(Figure [139]4E; Figure [140]SE3E, Supporting Information), size
distribution (Figure [141]4F; Figure [142]SE3F,G, Supporting
Information), or expression of EV marker proteins (CD63, CD9, and
TSG101) (Figure [143]4G; Figure [144]SE3H, Supporting Information)
between EVs from engineered cells and unmodified cells. The EVs derived
from the cells electro‐remolded with the IFN‐γ plasmid and PD‐L1 siRNA
exhibit significantly elevated levels of IFN‐γ mRNA (>6.52‐fold) and
PD‐L1 siRNA (>18.03‐fold) (Figure [145]4H,I; Figure [146]SE3I–L,
Supporting Information), demonstrating efficient packaging of
therapeutic molecules into SEE‐EVs. To further validate the modulatory
effects of SEE‐EVs on tumor cells, we incubated tumor cells with
SEE‐EVs and assessed IFN‐γ and PD‐L1 expression. The treated tumor
cells exhibited a significant increase of IFN‐γ mRNA (>2.27‐fold)
(Figure [147]4J; Figure [148]SE3M,N, Supporting Information) alongside
a marked reduction in PD‐L1 mRNA (>12.42‐fold) (Figure [149]4K; Figure
[150]SE3O,P, Supporting Information). Correspondingly, protein analysis
revealed elevated IFN‐γ (>2.71‐fold) and suppressed PD‐L1 expression
(>1.43‐fold) (Figure [151]4L–N; Figure [152]SE3Q–V, Supporting
Information).
Taken together, these results demonstrate that the procedure of
SEE‐mediated electro‐transfecting IFN‐γ plasmid and PD‐L1 siRNA into AG
cells, followed with efficient co‐loading of IFN‐γ mRNA and PD‐L1 siRNA
in SEE‐EVs. The engineered SEE‐EVs increase the IFN‐γ and downregulate
the PD‐L1 of targeted tumor cells.
2.5. The SEE‐EVs Mediated Metastasis Inhibition in CDX Model
To validate the capacity of SEE‐EVs in suppressing tumor metastasis in
vivo, we evaluated their therapeutic efficacy in the CDX model.
Corresponding to clinical situations, two application scenarios were
established by the CDX model, including metastasis prevention and
metastasis treatment (see details in Experimental Section). Metastasis
prevention refers to pharmacotherapy post‐resection to prevent
metastasis originating from incompletely resected tumor remnants or
circulating tumor cells, primarily applied to patients deemed eligible
for surgical tumor resection. Metastasis treatment refers to direct
drug administration for patients diagnosed with pre‐existing metastatic
tumors.^[ [153]^1 , [154]^9 ^]
In the CDX metastasis prevention scenario, mice were divided into five
groups (n = 6) according to the therapeutic agents: Control (PBS), EV
(EVs derived from untreated AG cells), EV@IFN‐γ (EVs derived from AG
cells loaded with IFN‐γ mRNA), EV@PD‐L1 (EVs derived from AG cells
loaded with PD‐L1 siRNA), and EV@IFN‐γ+PD‐L1 (EVs derived from AG cells
loaded with both the IFN‐γ mRNA and PD‐L1 siRNA) (Figure [155]5A). The
results show that the EVs co‐modified with IFN‐γ mRNA and PD‐L1 siRNA
significantly improve survival and tumor suppression. The single‐factor
engineered EVs exhibit moderate therapeutic effects, and the EV group
shows no differences compared to the control group (Figure [156]5B–D;
Figure [157]S12A–G, Supporting Information).
Figure 5.
Figure 5
[158]Open in a new tab
Tumor metastasis inhibition by SEE‐EVs in CDX model. A) The
construction and therapeutic regimen of CDX model mice with a risk of
tumor metastasis (n = 6 mice per group). B) Survival curves of CDX
model mice after different treatments. C,D) Growth curves of tumor
volume (C) and tumor to body weight ratios (D) of different treatment
groups. E) Photographs of lung tissue after different treatments. The
purple circles indicate the location of metastatic nodules. F) Number
of lung metastatic nodules detected in each group. G) H&E (top), Ki67
(middle), and TUNEL staining (bottom) of lung tissues (scale bars =
100 µm). The white circles indicate TUNEL‐positive cells. H,I)
Percentage of Ki67 (H) and TUNEL (I) positive cells among all cells
after the treatment ended in each group. J) Images of H&E staining of
liver tissue sections after different treatments (scale bars = 100 µm).
The black arrows indicate more aggregated abnormal cells. K) Number of
abnormal cells in liver tissue after different treatment. L) Phenotypic
proportion of peripheral blood macrophages in different treatment
groups. M,N) Percentage of CD4^+ T cells (M) and CD8^+ T cells (N) in
different treatment groups. The data were derived from three
independent experiments unless otherwise stated and were presented as
mean ± SD; a two‐sided Student's t‐test was used for comparisons (D, F,
H, I and K–N). p < 0.05 was considered statistically significant.
To further evaluate EV‐induced inhibition of tumor metastasis,
pulmonary metastatic burden was analyzed. According to lung nodules,
the EV@IFN‐γ+PD‐L1 group leads to the lowest number among all groups
(Figure [159]5E,F). In addition, the lowest cell proliferation rate and
the highest apoptosis rate are demonstrated in this group by the
histopathological assays such as H&E staining, Ki67 (proliferation
marker), and TUNEL (cell apoptosis) (Figure [160]5G–I). Assessment of
extrapulmonary metastasis reveals that the EV@IFN‐γ+PD‐L1 group
displays no visible metastatic lesions through H&E staining, while the
EV@IFN‐γ and EV@PD‐L1 groups show a small number of mitigated lesions
in the liver. By contrast, the control/EV groups exhibit prominent
hepatic metastases, accompanied by abnormal hepatic function indicators
(AST and ALB) (Figure [161]5J,K; Figure [162]S13A–I, Supporting
Information). To investigate the reasons for the variations in
treatment outcomes, we analyzed the therapeutic pathways, including
macrophage polarization and T cell activation in peripheral blood. The
results indicate that the EV@IFN‐γ+PD‐L1 group exhibits the most
pronounced shift toward M1 macrophages and the highest frequency of
activated T cells (Figure [163]5L–N; Figure [164]S14A–C, Supporting
Information).
For the metastasis treatment scenario, mice were randomly divided into
five groups receiving the same treatment regimens as in the metastasis
prevention scenario (Figure [165]SE4A, Supporting Information).
Following treatment, the engineered EVs (EV@IFN‐γ, EV@PD‐L1, and
EV@IFN‐γ+PD‐L1) lead to significantly smaller volumes of tumors as
compared to the control and EV groups. EV@IFN‐γ+PD‐L1 group results in
the most pronounced reduction (Figure [166]SE4B–J, Supporting
Information). The histopathological analysis further reveals the lowest
proliferation rate and the most apoptotic cells in the EV@IFN‐γ+PD‐L1
group (Figure [167]SE4K–M, Supporting Information). Further, we
investigated changes of macrophage subtypes and T cell activation
within tumor tissues. Among all groups, the EV@IFN‐γ+PD‐L1 group
demonstrates the highest M1 macrophage polarization and the most T cell
activation, indicating immune activation driven by the dual therapeutic
molecules encapsulated in EV@IFN‐γ+PD‐L1 (Figures [168]SE4N–P and
[169]S15A–C, Supporting Information).
Furthermore, we evaluated the biosafety of SEE‐EVs for tumor therapy.
H&E staining of major organs (heart, liver, spleen, lungs, kidneys)
reveals no histopathological abnormalities or inflammatory infiltration
in mice treated with SEE‐EVs (Figure [170]S16A, Supporting
Information). Serum biochemical analysis further confirms the absence
of hepatotoxicity or nephrotoxicity, with all liver (e.g., AST) and
kidney function markers (e.g., BUN) within normal ranges (Figure
[171]S16B–I, Supporting Information).
Taken together, the SEE‐EVs demonstrate biocompatibility and safety
profile in vivo, suggesting broad applicability for drug delivery.
Based on the findings that AG cells orchestrate their immune escape by
coordinately inhibiting macrophage and T cell activities, the SEE‐EVs
engineered with IFN‐γ mRNA and PD‐L1 siRNA exhibit enhanced therapeutic
performance in metastasis prevention and cancer treatment.
2.6. The SEE‐EVs Mediated Metastasis Inhibition in PDX Model
Based on the validation in the CDX model, we further conducted in vivo
application of the SEE‐EVs for metastasis inhibition in the PDX model,
a clinically relevant model that recapitulates tumor heterogeneity and
stromal microenvironment of patients.^[ [172]^36 , [173]^37 , [174]^38
, [175]^39 ^] Histomorphology profile and serum biochemical analysis
first confirmed the in vivo biosafety of the SEE‐EVs (Figure
[176]S17A–I, Supporting Information). To further validate the efficacy
of the SEE‐EVs in metastasis inhibition, both application scenarios of
metastasis prevention and treatment were established.
For the metastasis prevention scenario, a portion of tumor tissue was
cultured in vitro, followed by AG cell remodeling and SEE‐EVs
engineering. The remaining tissue was used to construct the PDX model
(Figure [177]6A; Figure [178]S18A–D, Supporting Information). Upon
completion of the model, SEE‐EVs were administered to the mice
according to their respective groups for therapeutic efficacy
evaluation. Compared to the other four groups, the EV@IFN‐γ+PD‐L1 group
demonstrates the highest rate of suppression to tumor growth and the
lowest number of metastatic pulmonary nodules (Figure [179]6B–E; Figure
[180]S19A–G, Supporting Information). Additionally, histopathological
assays indicate that the EV@IFN‐γ+PD‐L1 group displays optimal
pathological outcomes, with the highest efficacy in inhibiting tumor
cell proliferation and promoting apoptosis (Figure [181]6F–H).
Assessment of extrapulmonary metastasis reveals no detectable
metastases in the EV@IFN‐γ+PD‐L1 group, a low number of lesions in the
EV@IFN‐γ and EV@PD‐L1 groups, and a significantly higher number of
metastatic lesions in the control and EV groups (Figure [182]6I–J;
Figure [183]S20A, Supporting Information). Serum biochemical analysis
also reveals abnormal markers (AST, ALT, and ALB) exceeding the normal
threshold in the control and EV groups, whereas the other three
treatment groups maintain normal hepatic profiles (Figure [184]S20B–I,
Supporting Information). These results validated the therapeutic
efficacy of EV@IFN‐γ+PD‐L1 in preventing metastasis. Further analysis
demonstrates the synergistic promotion of M1 macrophage polarization
and T cell activation in EV@IFN‐γ+PD‐L1 group, which improves its
efficacy in tumor elimination (Figure [185]6K–M; Figure [186]S21A–C,
Supporting Information).
Figure 6.
Figure 6
[187]Open in a new tab
Inhibition of tumor metastasis by SEE‐EVs in PDX model. A) Construction
and therapeutic regimen of PDX model mice with a risk of tumor
metastasis (n = 6 mice per group). B,C) Growth curves of tumor volume
(B) and the final weight of tumors (C) in different treatment groups.
D) Photographs of lung tissue after different treatment. The purple
circles indicate the location of metastatic nodules. E) Number of lung
metastatic nodules detected in each group. F) H&E (top), Ki67 (middle),
and TUNEL staining (bottom) of lung tissues (scale bars = 100 µm). The
white circles indicate the location of TUNEL‐positive cells. G,H)
Percentage of Ki67 (G) and TUNEL (H) positive cells among all cells at
the end of treatment in each group. I) H&E staining of liver tissues
after different treatments (scale bars = 100 µm). The black arrows
indicate large aggregations of abnormal cells. J) Number of abnormal
cells in liver tissues after different treatments. K) Phenotypic
polarization of peripheral blood macrophages in different treatment
groups. L,M) Percentage of CD4^+ T cells (L) and CD8^+ T cells (M) in
different treatment groups. The data were derived from three
independent experiments unless otherwise stated and were presented as
mean ± SD; a two‐sided Student's t‐test was used for comparisons (C, E,
G, H, and J–M). p < 0.05 was considered statistically significant.
Subsequently, we constructed the metastasis treatment scenario for
evaluating therapeutic efficacy (Figures [188]SE5A and [189]S22A–D,
Supporting Information). The EV@IFN‐γ+PD‐L1 group demonstrates
significant suppression of tumor growth among all groups (Figure
[190]SE5B–J, Supporting Information). Histopathological evaluation
reveals no notable differences between the EV and control groups. The
EV@IFN‐γ+PD‐L1 group results in the lowest proliferation rate and
highest apoptotic activity in tumor cells (Figure [191]SE5K–M,
Supporting Information). Further cell analysis reveals the synergetic
activation of the immune system through M1 macrophage polarization and
T cell‐mediated cytotoxicity, which achieves significantly improved
metastasis treatment (Figures [192]SE5N–P and [193]S23A–C, Supporting
Information).
Taken together, the application of SEE‐EVs in PDX models demonstrates
its adaptability in clinical mimetic systems. Furthermore, these
results demonstrate the advantages of synergetic activation of
macrophage and T cells in tumor metastasis inhibition.
3. Discussion
In this work, we developed a nano‐electro‐platform addressing the
challenge in cancer therapy‐tumor metastasis inhibition. The platform
employs an evolutionary screening module for non‐destructive screening
the aggressive cell subpopulation. The platform further implements a
unique nanopore‐focused electric field, achieving efficient delivery of
therapeutic molecules into the AG cells, while triggering the remolded
cells to release EVs with enhanced tumor‐targeting and therapeutic
properties.
The SEE platform offers distinct advantages over existing methods. 1)
For isolating aggressive cell subpopulation, current methods, such as
flow cytometry^[ [194]^6 ^] and optical tweezers,^[ [195]^7 ^] face
challenges due to the lack of multiple identification channels for
precise cell profiling and screening. The SEE platform, incorporating
gradient culture zones, enables cell screening in terms of cellular
invasiveness, which bypasses the reliance on biomarkers in conventional
approaches. The successful isolation on the SEE platform and molecular
characterization of highly invasive cell subpopulations yield
mechanistic insights into tumor metastasis, while offering a foundation
for developing targeted therapeutic strategies. Considering the wide
applicability of the SEE platform for aggressive cell screening, other
cells can be expanded for further studies, such as hepatocellular
carcinoma cells, a high‐incidence cancer type. 2) Compared with other
exogenous carriers‐based drug delivery systems, EVs derived from
autologous cells (patient‐derived) minimize immune rejection risks,
ensuring biological compatibility.^[ [196]^40 , [197]^41 , [198]^42 ^]
Although tumor cell‐derived EVs have been reported to exhibit targeting
capabilities toward tumor in vivo,^[ [199]^19 , [200]^43 ^] in this
work, we considered the presence of cellular heterogeneity within
tumors while adding the note that the significantly enhanced
tumor‐targeting capability EVs derived from the aggressive cell
subpopulation. In addition, the inherent diversity of EVs suggests the
potential to directly isolate functionally distinct EV subsets from
heterogeneous populations for tailored therapeutic applications.^[
[201]^44 , [202]^45 ^] 3) Compared to traditional cell engineering
methods,^[ [203]^46 ^] the platform‐mediated electro‐remodeling of
cells for therapeutic EVs engineering, as evidenced by EVs co‐loaded
with IFN‐γ mRNA and PD‐L1 siRNA, presents a novel strategy that
overcomes the limitations of conventional non‐targeted drug‐based
therapies in suppressing metastatic progression. Given the versatility
of the SEE platform for molecular delivery, future improvements could
focus on expanding the range of deliverable molecules to enhance
therapeutic efficacy through multiple mechanisms.
4. Conclusion
Tumor metastasis is the leading cause of mortality in cancer patients,
making mechanism research and effective treatment imperative. Due to
the high heterogeneity of tumors, metastatic foci are predominantly
formed by a subpopulation of aggressive cells behaving enhanced
migration and invasiveness. Screening the aggressive subgroup for
decoding the underlying mechanism brings about important hint for
inhibiting tumor metastasis.
In this work, we developed a nano‐electro‐platform addressing the
challenge in cancer therapy‐tumor metastasis inhibition. The platform
employs an evolutionary screening module for non‐destructive screening
the aggressive cell subpopulation. The platform further implements a
unique nanopore‐focused electric field, achieving efficient delivery of
therapeutic molecules into the AG cells, while triggering the remolded
cells to release EVs with enhanced tumor‐targeting and therapeutic
properties that efficiently transport into targeted tumor cells. The
engineered EVs effectively activate macrophages and T cells, leading to
robust tumor cell elimination and metastasis inhibition in lung cancer
metastasis models. These highlight a versatile, multidisciplinary
technique adopting a new path toward deep understanding and treating
metastasis and overcome the limitations of conventional non‐targeted
drug‐based therapies in suppressing metastatic progression.
5. Experimental Section
Fabrication of SEE Platform
The dimensions and structures of each layer of the SEE platform are
illustrated in Figure [204]S1 (Supporting Information). Layer (1) was
fabricated using polydimethylsiloxane (PDMS, Dow Corning, SylgardTM
184, MI, USA), which was prepared by mixing the elastomer base with
curing agent at a 10:1 (w/w) ratio and degassing under vacuum
(‐0.1 MPa) for 30 min. After cured at 80 °C for 30 min, the PDMS layer
was peeled from the mold and perforated to form inlets and outlets.
Layers (2), (3), and (4) were crafted from acrylic plate utilizing
laser cutting technology. Layer (5) consisted of indium tin oxide (ITO)
glass with a resistance of 6 Ω (NOZO, China). Prior to assembly, layers
(1) and (2) were treated with oxygen plasma and bonded at 80 °C for
2 h. Layers (2), (3), (4), (5), and (6) were then assembled using
acrylic adhesive. Finally, the prepared chips were rinsed with 75%
ethanol and sterile water, followed by UV sterilization for 1 h.
Cell Culture
Human non‐small cell lung cancer A549 cells were cultured in F‐12K
medium (21127022, Gibco, USA) containing 10% fetal bovine serum (FBS)
(A5670701, Gibco, USA) and 1% penicillin‐streptomycin (G4015,
Servicebio, China). Human non‐small cell lung cancer H1975 cells, human
umbilical vein endothelial HUVEC cells, and mouse lung carcinomas Lewis
cells were cultured in RPMI 1640 medium (11875119, Gibco, USA)
containing 10% FBS and 1% penicillin‐streptomycin. ID8 epithelioid
cells and human hepatocellular carcinomas HepG2 cells were cultured in
DMEM medium (11965092, Gibco, USA) containing 10% FBS and 1%
penicillin‐streptomycin. All cell lines were maintained at 37 °C with
5% CO[2] in a cell incubator (ThermoFisher Scientific, USA).
Workflow of SEE Platform
A suspension of 2 × 10^6 tumor cells was introduced into the ES module
via the cell inlet. Following one week of incubation, AG cells migrated
upward along the sloped surface. Cell culture medium was then removed
and the chip was turned over. 0.05% trypsin solution (G4011,
Servicebio, China) was introduced through the cell inlet (not in
contact with the cells at the bottom) and incubated with the cells on
the slope for 5 min at room temperature. Upon complete digestion of the
AG cells, an EVs‐free culture medium was added via the buffer inlet to
terminate digestion and facilitate cell transfer into the ER module.
The chip was then reverted to its original orientation, enabling cell
attachment to the nanomembrane. After a 6 h period, the molecules to be
delivered (e.g., 5 µg IFN‐γ plasmid, 20 nm PD‐L1 siRNA (Table [205]S1,
Supporting Information) were added through the cargo inlet. A gold
needle was pierced into the first layer of the chip to serve as the top
electrode and connected to the positive electrode of an electroporator
(ECM830, BTX, USA), while the ITO bottom electrode was linked to the
negative terminal. Following pulsed electric field application, the top
electrode was removed. The cells were subsequently cultured for an
additional 48 h before supernatant collection for EVs extraction.
Cell Scratch Assay
≈2 × 10^6 cells were uniformly seeded into culture dishes. Upon
reaching 80–90% confluency, a straight scratch was introduced across
the cell monolayer using a sterile 200 µL pipette tip under a
microscope (CKX3‐SLP, OLYMPUS, Japan). Following PBS (G4202,
Servicebio, China) washes, the scratch location was marked on the dish
bottom for consistent tracking. Serum‐free medium was then added, and
the dish was returned to the incubator. Cell migration behavior was
monitored and documented at 0 (baseline), 24, 48, and 72 h
post‐scratching using phase‐contrast microscopy.
Cell Invasion Assay
The 3D collagen matrix was prepared using Mouse Tail Collagen Type I
Kit (200110‐10, Shengyou, China) following the manufacturer's protocol.
Briefly, 200 µL mouse tail collagen type I was mixed with 12 µL
0.1 mol L^−1 NaOH and 23 µL 10 × PBS (G4202, Servicebio, China). Then,
the mixture (100 µL) was evenly spread onto a confocal dish and
solidified at 37 °C for 20 min. Next, cells were stained with DAPI
(GDP1024, Servicebio, China) for 15 min and seeded onto the collagen
matrix for 30 min. The remaining collagen mixture was then overlaid
onto the cells. At designated time points, the cells were monitored
using a live‐cell imaging station on a fluorescence confocal microscope
(Dragonfly, Andor, UK). The rate of invasive cells was calculated as
the percentage of invasive cells relative to the total cells.^[
[206]^47 , [207]^48 ^] Invasive cell number was quantified as cells
located ± 200 µm vertically from the inoculation plane (Z = 0 µm)
(Figure [208]S5A, Supporting Information). The total cell number was
the sum of invasive cells and those remaining on the original
inoculation plane.
Collection and Purification of EVs
The cell supernatant containing EVs was sequentially centrifuged at 300
× g for 10 min to remove cells, followed by 2000 × g for 30 min to
eliminate cellular debris. The clarified supernatant was then
centrifuged at 10 000 × g for 30 min at 4 °C to remove micro‐vesicles.
Finally, EVs were isolated via ultracentrifugation at 110 000 × g for
90 min at 4 °C. The morphology of EVs were examined using a
transmission electron microscope (JEM‐1400, JEOL, Japan), while their
size distribution and concentration were determined by nanoparticle
tracking analysis (Zetaview‐PMX120‐Z, Particle Metrix, Meerbusch,
Germany).
Western Blot Analysis
Following the samples lysis with RIPA buffer (PC101, Epizyme Biotech,
China), protein concentrations were quantified using a BCA protein
assay kit (ZJ102, Epizyme Biotech, China). Then, the protein samples
were separated by 12.5% PAGE gel (PG113, Epizyme Biotech, China) and
subsequently transferred onto polyvinylidene fluoride (PVDF) membranes
(88518, ThermoFisher, USA). The membranes were blocked with 5% BSA
(PS113, Epizyme Biotech, China) in Tris‐buffered saline (TBS, PS103,
Epizyme Biotech, China) containing 0.1% tween 20 (9005‐64‐5, Beyotime,
China) and then incubated overnight at 4 °C with primary antibodies
(details are provided in Table [209]S2, Supporting Information). After
washing, the membranes were probed with HRP‐conjugated secondary
antibodies for 40 min at room temperature. Protein bands were
visualized using an enhanced chemiluminescence detection system (GE
Healthcare, UK).
RNA Extraction and RT‐qPCR Assay
Total RNA was extracted using a commercial kit (RC112, Vazyme, China)
following the manufacturer's instruction. Target gene expression was
analyzed by quantitative reverse transcription PCR (RT‐qPCR) using
HiScript II One Step qRT‐PCR SYBR Green Kit (Q221, Vazyme, China). The
sequence of all primers is provided in Table [210]S1 (Supporting
Information).
EV Internalization Analysis
EVs were labeled with PKH26 dye (MINI26‐1KT, Sigma, USA) for 10 min at
room temperature. The staining reaction was quenched by adding FBS,
followed by ultracentrifugation to isolate the PKH26‐labeled EVs. For
the uptake assay, 10 µg PKH26‐labeled EVs were co‐incubated with
different cells for 12 h. After incubation, unbound EVs were removed by
washing, and the cells were fixed with 4% paraformaldehyde (G1101,
Servicebio, China). Nuclei were counterstained with DAPI prior to
fluorescence imaging.
Liposome‐Mediated Delivery Assay
The lipofectamine (L3000015, ThermoFisher, USA) was used to deliver
plasmid (5 µg) into the cells (2 × 10^6) according to the recommended
ratio. After transfection for 6 h, the medium was replaced with a
complete medium, and the cells were cultured for an additional 24 h
before further analysis.
BEP‐Mediated Delivery Assay
[MATH: ≈ :MATH]
2 × 10^6 cells were mixed with 5 µg plasmid in 500 µL Opti‐MEM medium
(31 985 070, ThermoFisher, USA). Electroporation was performed using
the parameters of 30 V, 1 ms. Following electroporation, the cells were
incubated in a complete medium for 24 h prior to further analysis.
Cell Viability Assay
Cell viability was evaluated using the cell counting kit‐8 (CCK‐8)
assay. Pre‐treated cells (≈1 × 10^4 cells/well) were seeded into a
96‐well plate for 24 h. Subsequently, 10 µL CCK‐8 solution (CK18,
Dojindo, China) was added to each well, followed by incubation for
1–4 h at 37 °C. The absorbance at 450 nm was then measured using a
microplate reader (TECAN, Spark, Switzerland). Cell viability was
calculated as the ratio of the absorbance of the experimental group to
that of the control group.
Delivery Efficiency Analysis
Following plasmid delivery, the cells were detached using 0.25% trypsin
solution (G4013, Servicebio, China). The cell suspension was
centrifuged at 500 × g for 5 min and washed twice with PBS. Finally,
the cells were resuspended in 300 µL PBS and analyzed using a flow
cytometer (DxFLEX, Beckman Coulter). The delivery efficiency was
calculated as the percentage of GFP‐positive cells (expressing the
GFP‐labeled plasmid) relative to the total cell population.
Detection of EVs Yield
The total protein content of EVs secreted by an equal number of cells
(1.5 × 10^7) across all treatment groups was quantified using the BCA
protein assay kit following the manufacturer's protocol, serving as the
EV production for each treatment.
Collection and Processing of Clinical Tumor Tissue Samples
Lung cancer tissue specimens were provided by Peking University Cancer
Hospital & Institute. This study was approved by the Ethics Committee
of Peking University Cancer Hospital & Institute (Institutional Review
Board No. 2023KT146). The information of clinical patients that
provided tumor tissue samples in Table [211]S3 (Supporting
Information). All sample donors signed informed consent forms. The
excised tissues were preserved in a complete medium containing 10% FBS.
After PBS rinsing to remove blood and impurities, the specimens were
mechanically dissociated into small fragments (≈1 mm^3 in diameter).
These fragments were then digested in a 3 mL digestion solution,
consisting of 2% FBS, 1 mg mL^−1 collagenase I (C8140, Solarbio,
China), and 200 µg mL^−1 DNase I (D8071, Solarbio, China), at 37 °C for
2 h. The resulting suspension was filtered through a 70 µm cell
strainer (352350, Corning, USA) and centrifuged at 350 × g for 10 min.
Following PBS washing and repeat centrifugation, the cell pellet was
resuspended in F‐12K medium supplemented with 15% FBS and FibrOut
fibroblast inhibitor (4‐21564, CHI Scientific, China) in a cell
incubator at 37 °C with 5% CO[2].
Collection and Processing of Clinical Blood Samples
Clinical blood samples were placed in blood collection tubes containing
anticoagulants and diluted with PBS (blood: PBS = 1:1). Peripheral
blood mononuclear cells (PBMCs) were isolated using human whole blood
mononuclear cell separation solution (P9011, Solarbio, China). Briefly,
the diluted blood was layered onto an equal volume of separation medium
(half the volume of the diluted blood) and centrifuged at 800 × g for
20 min, with acceleration and deceleration rates set to 1. After
centrifugation and washing with PBS, the PBMCs were preserved with
cryopreservation solution (G1709, Servicebio, China) in liquid
nitrogen.
Animal Experiments
Female mice, including BALB/c nude (5–8 weeks), BALB/c (5–8 weeks),
C57BL/6 (5–8 weeks), and NTG (4–5 weeks) mice, were obtained from SPF
Laboratory Animal Technology in Beijing, China (Animal Certification
No. SCXK 2019‐0010). Animals were kept in isolator cages within a
pathogen‐free facility. All animal experiments were approved by the
Laboratory Animal Ethics Committee of Beijing Cancer Hospital
(Institutional Review Board No. 2023KT146) and conducted in compliance
with the Guidelines on Humane Treatment of Laboratory Animals and
standard procedures for animal care and use.
In Vivo biodistribution of EVs
[MATH: ≈ :MATH]
5 × 10^6 tumor cells were subcutaneously injected into the flanks of
BALB/c nude or BALB/c mice. One week later, EVs derived from tumor
cells were labeled with DiD (V22887, ThermoFisher, USA) for 20 min and
injected into mice via the tail vein. In vivo fluorescence imaging was
performed using the IVIS Spectrum imaging system (Revvity, USA).
Following imaging, the mice were euthanized, and tumors along with
major organs were harvested for ex vivo imaging and frozen section.
Tumor Metastasis Prevention Assay
In this study, both CDX and PDX tumor metastasis models were
established. For the CDX model, BALB/c mice received subcutaneous
injections of 5 × 10^6 Lewis cells into the flank region. For the PDX
model, NTG mice underwent surgical implantation of patient tumor
fragments (2–3 mm^3) into the inguinal region using hookless ophthalmic
tweezers. Following two generations of tumor expansion (F1 and F2), the
third generation (F3) tumor tissues were re‐transplanted into new NTG
mice. Two weeks post‐engraftment, 7 × 10^6 PBMCs were injected
intraperitoneally to generate humanized mice with reconstituted immune
systems. Immune reconstitution was verified through flow cytometric
analysis of human CD45^+ T cells in orbital blood. Meanwhile,
Graft‐versus‐host disease (GVHD) was monitored twice weekly through
clinical scores (based on weight, posture, activity, hair coat, skin,
etc.). In both models, when tumors reached 40 mm^3, visible tumor
tissue was surgically excised and 2 × 10^5 tumor cells were injected
intravenously via the tail vein to simulate the scenario of tumor
metastasis in vivo. Mice were then randomized into five treatment
groups receiving different EVs preparations (100 µg per injection) via
tail vein every two days for five total administrations. Following the
treatment regimen, mice were euthanized for collection of blood, tumor
tissues, and major organs to evaluate therapeutic efficacy.
Tumor Metastasis Treatment Assay
[MATH: ≈ :MATH]
5 × 10^6 aggressive cells, selected through evolutionary screening on
the SEE platform, were subcutaneously injected into BALB/c mice (CDX
model) or NTG mice (PDX model). When tumors reached 80–100 mm^3, mice
were randomized into five treatment groups receiving different EVs
preparations (100 µg per injection) via tail vein every two days for
five total administrations. Following the treatment regimen, mice were
euthanized for collection of blood, tumor tissues, and major organs to
evaluate therapeutic efficacy.
Histology and Immunohistochemistry Analysis
Tissue samples including heart, liver, spleen, lung, kidney, and tumor
were fixed in 4% paraformaldehyde overnight, followed by paraffin
embedding and frozen section at 10 µm thickness. For histological
processing, all paraffin sections underwent dewaxing and hydration
through sequential immersion in: 1) environmentally friendly dewaxing
solution (G1128, Servicebio, China) I 10 min, 2) environmentally
friendly dewaxing solution II 10 min, 3) environmentally friendly
dewaxing solution III 10 min, 4) anhydrous ethanol I 5 min, 5)
anhydrous ethanol II 5 min, and 6) anhydrous ethanol III 5 min, with a
final rinse in tap water. Subsequently, the sections were subjected to
hematoxylin‐eosin (HE) staining, Ki67 immunohistochemistry, and TUNEL
apoptosis assay.
HE Staining
HE staining was performed on dewaxed sections using a hematoxylin‐eosin
staining kit (G1076, Servicebio, China) according to the manufacturer's
protocol. The sections underwent sequential steps, including dewaxing,
staining pretreatment, hematoxylin staining, eosin staining,
dehydration, and sealing. After processing, images were acquired and
analyzed, with nuclei staining blue and cytoplasm staining red.
Ki67 Staining
Dewaxed sections were processed through the following sequential steps:
antigen recovery, endogenous peroxidase blocking, non‐specific site
blocking, primary antibody incubation ([212]GB121141, Servicebio,
China), and secondary antibody incubation (GB23301, Servicebio, China).
Following PBS washes, sections were treated with freshly prepared DAB
chromogenic solution (G1212, Servicebio, China). The reaction was
terminated by rinsing with tap water. Subsequently, sections were
counterstained with hematoxylin, dehydrated, and mounted.
Immunostaining results were evaluated under microscopy.
TUNEL Assay
Dewaxed sections were processed using a TUNEL staining kit (G1504,
Servicebio, China) following the manufacturer's protocol. Briefly,
slides were sequentially treated with protease K for antigen retrieval,
permeabilized, equilibrated at room temperature, and incubated with the
reaction mixture (TDT enzyme: dUTP: buffer = 1: 5: 50). After nuclear
counterstaining with DAPI, the sections were mounted for fluorescence
microscopy imaging.
Kidney and Liver Function Analysis
Mouse peripheral blood was collected via orbital bleeding. Following a
2 h standing at room temperature, the samples were centrifuged at 1500
× g for 15 min. The supernatant was subsequently analyzed using an
automated biochemical analyzer.
Flow Cytometry Assay
The cells were suspended in 100 µL cell staining buffer (E‐CK‐A107,
Elabscience, China) and incubated with an anti‐CD16 antibody for
10 min. Subsequently, specific antibodies for T cells and macrophages
(details are provided in Table [213]S4, Supporting Information) were
added for cell identification. Following 30 min incubation, the cells
were added into 1 mL cell staining buffer and centrifuged at 350 × g
for 10 min at 4 °C. The cells were then resuspended in a fixation
buffer (E‐CK‐A109, Elabscience, China) for 30 min. After washes with
cell staining buffer, the cells were analyzed using flow cytometry
(LSRFORTESSA, Becton, Dickinson and Company, America).
Statistical Analysis
To simulate the distribution of the electric field, a simplified model
of the electro‐remodeling module was established by using COMSOL
Multiphysics (version 6.1). The data were analyzed using GraphPad Prism
8.0, Origin 8.0, Flowjo 10.8, and Image J FIJI software. Student's
t‐test and one‐way ANOVA were used for comparisons between groups. The
data were presented as mean ± standard deviation (SD). Differences were
considered statistically significant when p < 0.05.
Conflict of Interest
The authors declare no conflict of interest.
Supporting information
Supporting Information
[214]ADVS-12-e07684-s001.docx^ (11.3MB, docx)
Acknowledgements